(B-66) Nanoemulsions for Encapsulation of Bacteriophages Targeting Escherichia coli

Antimicrobial resistance (AMR) is a severe public health crisis worldwide affecting treatment options for a range of infections. It was associated with approximately 5 million deaths in 2019 alone.¹ AMR challenges the use of traditional antibiotics and raises the necessity of developing new antimicrobial agents.^2,3

Lytic bacteriophages have potential to ease this crisis. As highly abundant viruses, bacteriophages use a bacterial cell’s own mechanisms to replicate before destroying it. Some bacteriophages possess the ability to only target specific bacterial strains, while others possess the ability to target a broader spectrum of bacteria.⁴

Though bacteriophages carry several benefits as natural killers of bacteria, their therapeutic potential in living organisms can be inhibited due to issues involving stability and harsh conditions. Despite these shortcomings, systems on the nanoscale, such as nanoemulsions (NEs), have potential to encapsulate bacteriophages and better control their delivery to sites of infection.^5-7 Previously, bacteriophages targeting Staphylococcus aureus and Pseudomonas aeruginosa were incorporated in oil-in-water (O/W) emulsions and water-in-oil-in-water (W/O/W) multiple emulsions respectively, where they demonstrated noticeable antibacterial effects.^8,9

This proof-of-concept study discusses the incorporation of bacteriophages into the aqueous phase of a water-in-oil (W/O) NE to target Escherichia coli (E. coli). NEs and bacteriophages were characterized to evaluate their feasibility. The aqueous phase weight percentages and final emulsification methods of the NEs were varied. Traditional double layer plaque assays (DLPAs) and zone of inhibition (ZoI) studies were conducted to assess differences in antibacterial efficacy.

Bacteriophages were grown by inoculating a log phase E. coli liquid culture with thawed bacteriophage stock. The culture underwent centrifugation once maximum bacteriophage growth was reached. To enumerate bacteriophages, 5 mL soft agar, 300 µL E. coli, and 100 µL diluted culture supernatant were combined, poured over Luria-Bertani (LB) agar plates, and incubated overnight. Plaques were counted to determine bacteriophage titer in plaque forming units (PFU).

NEs were fabricated on a stir plate at 700 rpm. Injections of 200 µL vacuum-filtered (0.1-micron, 47 mm PTFE laminated membrane filter) oily phase (70 parts/weight hexadecane, 22.5 parts/weight Span 80, and 7.5 parts/weight Tween 80) were added to the aqueous phase at 20 s intervals. Afterward, NEs remained stirring for 1–2 h or received ultrasonication for 2.5 mins (50% power, approximately 19.5 kHz). NEs containing LB and bacteriophages were handled aseptically (excluding ultrasonication), remade before each new set of experiments, stored at RT, and mixed before use.

Dynamic light scattering (DLS) (Zetasizer Nano ZS by Malvern Panalytical) characterized NEs. For bacteriophage size quantification via transmission electron microscopy (TEM) (JEOL JEM 1011), bacteriophages were ultracentrifuged, resuspended in DI H₂O, and stained with 1% uranyl acetate for 2 mins. For DLPAs, 5 mL soft agar, 100 µL E. coli, and 100 µL NE were combined and poured over LB agar plates. For ZoI studies, 50 µL E. coli was spread over LB agar plates. The NEs were pipetted directly onto UV-sterilized 6 mm paper disks placed atop the inoculated plates. ImageJ quantified zone areas.

For DLS, DI H₂O was used as the aqueous phase. Additionally, NEs underwent dilutions (1:100) in hexadecane. The lower polydispersity indexes (PDIs) of the 10 v/v% aqueous phase NEs represent more homogenous droplet size distributions than in the 5 v/v% aqueous phase NEs. Additionally, with these NEs, the emulsification method produced a larger difference in droplet size (94.9 ± 0.82 nm with stirring vs. 52.9 ± 1.97 nm with ultrasonication). However, all NEs contained nanometer-sized droplets. From TEM imaging, bacteriophages had an average height of 64.01 ± 2.5 nm, so bacteriophages in most NEs were likely not fully encapsulated.

Both DLPAs and ZoI indicated NEs with bacteriophages (titer about 1.65 x 10⁹ PFU) hindered E. coli growth, while NEs without bacteriophages did not. This suggests that only active bacteriophages, not the oily phase, were bactericidal. In ZoI, noticeable zones were present with the NEs containing bacteriophages. Interestingly, the NE fabrication method that produced the smallest average droplet size (5 v/v% aqueous content, stirring) formed the largest zones (630.61 ± 238.03 mm²). Additionally, the NE fabrication method that produced the largest average droplet size (10 v/v% aqueous content, stirring) generated the smallest zones (82.16 ± 43.83 mm²).

In larger droplets, bacteriophages were more likely to be fully encapsulated and take longer to release, while in smaller droplets, bacteriophages were likely only partially encapsulated and dispersed faster. Therefore, correlation studies between droplet sizes and time of bacteriophage release are necessary to control their delivery and evaluate their full therapeutic potential. Additionally, since both NEs containing 10 v/v% bacteriophages had smaller zones than the NEs containing 5 v/v% bacteriophages, the effects of aqueous phase weight percentage on timing of bacteriophage release should continue to be explored.

We successfully incorporated bacteriophages into NEs and demonstrated their ability to inhibit E. coli growth. As these NEs may be used to treat surgical site infections, they must be evaluated for mammalian cytotoxicity. If NEs show toxicity towards mammalian cells, natural base oils can possibly be used as alternatives to hexadecane in the oily phase. Ex vivo permeability and stability studies will help determine clinical utility.

This material is based upon work supported by a National Science Foundation Research Experiences for Undergraduates (REU) site program under Grant No. 1950581. James Geeter and Daphne Little provided sewage samples from which bacteriophages were isolated and purified by Dr. Vijay Singh Gondil. We would like to thank Dr. Artur Muszynski and the Complex Carbohydrate Research Center (CCRC) at the University of Georgia, as well as Mary Ard and Georgia Electron Microscopy for assistance with TEM preparation (ultracentrifugation) and imaging, respectively. TEM and ZoI images were analyzed in ImageJ. This research was conducted with the support of Drs. Hitesh Handa and Elizabeth J. Brisbois at the University of Georgia. The presenter would like to thank Sarah N. Wilson for her day-to-day mentorship during the REU program.

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